Low Po₂ conditions induce reactive oxygen species formation during contractions in single skeletal muscle fibers

Abstract

Contractions in whole skeletal muscle during hypoxia are known to generate reactive oxygen species (ROS); however, identification of real-time ROS formation within isolated single skeletal muscle fibers has been challenging. Consequently, there is no convincing evidence showing increased ROS production in intact contracting fibers under low Po₂ conditions. Therefore, we hypothesized that intracellular ROS generation in single contracting skeletal myofibers increases during low Po₂ compared with a value approximating normal resting Po₂. Dihydrofluorescein was loaded into single frog (Xenopus) fibers, and fluorescence was used to monitor ROS using confocal microscopy. Myofibers were exposed to two maximal tetanic contractile periods (1 contraction/3 s for 2 min, separated by a 60-min rest period), each consisting of one of the following treatments: high Po₂ (30 Torr), low Po₂ (3–5 Torr), high Po₂ with ebselen (antioxidant), or low Po₂ with ebselen. Ebselen (10 μM) was administered before the designated contractile period. ROS formation during low Po₂ treatment was greater than during high Po₂ treatment, and ebselen decreased ROS generation in both low- and high-Po₂ conditions (P < 0.05). ROS accumulated at a faster rate in low vs. high Po₂. Force was reduced >30% for each condition except low Po₂ with ebselen, which only decreased ∼15%. We concluded that single myofibers under low Po₂ conditions develop accelerated and more oxidative stress than at Po₂ = 30 Torr (normal human resting Po₂). Ebselen decreases ROS formation in both low and high Po₂, but only mitigates skeletal muscle fatigue during reduced Po₂ conditions.

reactive oxygen species (ros) play important roles in biological systems (1, 44, 46, 47, 49, 50). ROS have been documented as a general response to ischemia-reperfusion injury (26, 43), muscle stimulation (29), and heat stress (44). Excessive ROS disrupt nearly all physiological systems (1, 44, 46, 47, 49, 50). However, the underlying mechanism of ROS formation in hypoxic skeletal muscle has not been fully elucidated. Moreover, there has been little investigation of ROS generation within single skeletal muscle fibers under low Po₂ conditions using real-time measurements, which eliminates some of the problems associated with these measurements that have been made in whole animal or whole muscle preparations (46). Previous research suggests that in human skeletal muscle, intracellular Po₂ drops from ∼30 Torr at rest to 3–5 Torr during exercise (40). Thus, it is of interest to investigate intracellular ROS formation during these conditions of low Po₂ in single contracting myocytes.

Hypoxia causes a significant accumulation of reducing agents in the mitochondria, such as NADH and FADH₂. Abrupt exposure to O₂ can promote immediate formation of superoxide anion (O₂^·−) in the mitochondrial electron transport chain (2, 37), thus initiating oxidative stress. Moreover, substantial data point to intracellular ROS formation in cardiac tissue during hypoxic or ischemic conditions (6, 38). Low Po₂ environments have potent effects on nearly all biological systems. However, they may have particular relevance to skeletal muscle because myofibers consume large amounts of O₂ to function at optimal levels, as well as to replenish ATP hydrolyzed during contractions. Therefore, these muscles may undergo severe oxidative stress during hypoxia (Po₂ 3–5 Torr) (46).

In the present study, we tested the hypothesis that exposure to hypoxic conditions increases ROS production in intact contracting single frog (Xenopus) fibers. We also tested whether antioxidant administration abolishes the elevated levels of ROS produced in these low Po₂ conditions. Our results provide real-time insight into the molecular mechanisms of fatigue in an intracellular hypoxic environment, which likely involves ROS signaling and antioxidant defense systems.

MATERIALS AND METHODS

All procedures were approved by University of California, San Diego (UCSD), Institutional Animal Care and Use Committee (IACUC). Adult female Xenopus laevis were euthanized, lumbrical muscles (II–IV) from the foot were removed, and intact single myofibers were microdissected. Fast-twitch, glycolytic fibers were selected for use in the current study, as described previously (34). After dissection, the tendons of each myofiber were secured with platinum clips and attached in a glass-bottomed chamber to a force transducer (1500A Small Intact Muscle Test System and force transducer model 400A, Aurora Scientific). The chamber was loaded with Ringer's solution (in mM: 116.5 NaCl, 2 KCl, 1.9 CaCl₂, 2 NaH₂PO₄, and 0.1 EGTA, at pH 7.0) at 20°C. Each fiber was treated with 5 μM dihydrofluorescein diacetate (Hfluor-DA; Sigma), a fluorescent probe that is used for the detection of intracellular ROS, for 30 min. Ebselen (10 μM; Alexis Biochemicals) was incubated simultaneously with Hfluor-DA, and kept in the bath throughout the contractile period. A laser scan confocal microscope (McBain Systems with a Nikon inverted microscope) recorded fluorescent signals from the interior sections of the fibers. Fiber length was adjusted to produce maximal tetanic force and during the contractile period, each fiber was electrically stimulated every 3 s for 2 min at 20°C (S48 stimulator; Grass Technologies; 250-ms trains, 2-ms pulse duration, 70 Hz, 8 V). Each individual fiber had two separate contractile periods, which were separated by a 60-min rest period. The contractile periods for each fiber consisted of exposure to low Po₂ (3–5 Torr; n = 10) or a value that approximates resting normal human skeletal muscle Po₂ (30 Torr; n = 10); or these two Po₂ conditions with ebselen (n = 3). The treatment order was randomized to ensure that the order of intracellular Po₂ conditions did not influence the change in fluorescence. Hereafter, 30 Torr is referred to as “high” Po₂ while 3–5 Torr is referred to as “low” Po₂.

All confocal experiments were performed in a dark room. The parameters for confocal imaging setup for ROS measurement were listed as the following: laser, argon; excitation, 488 nm; and emission, 525 ± 15 nm. The background fluorescence was kept minimal during the experiments. An image (512 × 512 pixels) was captured every 5 s (to prevent photobleaching) with a 300-ms scanning time in a gated system, and the mean fluorescence of each image was used for the calculation of ROS (H₂O₂) formation.

Muscle contractile function and ROS formation during high Po₂ and low Po₂ conditions were measured simultaneously in each fiber. To overcome motion artifact, we followed a rigorous criterion with all of the fibers. When the fiber motion exceeded 10% of the size of the field in all directions based on visual landmark recognition, the data were discarded. We used Hfluor because it is more resistant to photo damage compared with its analogs (10, 46). To further reduce photo bleaching, we did not use a continuous mode of measurement, but selected a shutter program that opened the laser once for 300 ms every 5 s to capture each image. Moreover, the laser power was adjusted to a relatively low level without any significant attenuation of sensitivity or resolution.

Prior to the experiments conducted on single myofibers, we did preliminary experiments to test whether different oxygen tensions affect the reaction between Hfluor and hydrogen peroxide (H₂O₂). We found that that there was no significant difference among conditions of 3–5 Torr, 30 Torr, and 160 Torr, which agrees with former research, suggesting that oxygen tension is not a major factor for the reaction (10, 45). Furthermore, our experiments were performed at ∼20°C for the following reasons: 1) Xenopus frogs were housed in a water tank of ∼20°C in an animal facility. This temperature setting was approved by UCSD IACUC. Thus, this set-point is more consistent with normal physiological temperatures in amphibians. 2) It has been tested that the integrity and function of isolated myofibers are maintained longer at 20°C compared with 37°C or 10°C (data not shown), which is consistent with previous studies using 20°C as the ideal temperature when undertaking experiments using amphibian muscle fibers (11, 12, 34, 46).

Data were analyzed using a multi-way ANOVA and expressed as means ± SE (JMP; SAS Institute). The differences between the four treatments were identified by post-ANOVA contrast analysis from SAS JMP software. P < 0.05 was considered to be significant.

RESULTS

Representative images from Hfluor-loaded fibers are shown in Fig. 1. In Fig. 1, A and B, under confocal microscopy, ROS fluorescence increased after a 2-min tetanic contractile period under Po₂ 30 Torr. Likewise, fluorescence was enhanced after a contraction period under Po₂ 3–5 Torr (Fig. 1, C and D). However, this increase in fluorescence was greater than that of the Po₂ 30 Torr condition. Ebselen treatment completely abolished the increase in fluorescent signal during a 2-min contraction period in both Po₂ 30 Torr and Po₂ 3–5 Torr conditions (Fig. 1, E–H).

Fig. 1.

Representative images from dihydrofluorescein diacetate (Hfluor) loaded Xenopus single fibers. A: fiber before contractions. B: same area of image A after contractions with the perfusate at Po₂ 30 Torr (normal human muscle resting Po₂). C: a fiber before contractions. D: same area of image in C after contractions with the pefusate at Po₂ 3–5 Torr (low Po₂). E: fiber treated with ebselen before contractions. F: same area of image in E after contractions under Po₂ 30 Torr. G: a fiber treated with ebselen before contractions. H: same area of image in G after contractions under Po₂ 3–5 Torr.

Figure 2, A and B, illustrates typical ROS fluorescence during two 2-min tetanic contractile periods separated by a 60-min rest period. The experiment was conducted in a blocked order (i.e., 50% of fibers used were tested under high Po₂ in the first contractile bout followed by a second bout under low Po₂, and the other 50% were tested in the reverse sequence) to ensure that the order of intracellular Po₂ conditions did not affect the change in fluorescent intensity. This figure demonstrates that the contractile period elicited less fluorescence when the Po₂ was high.

Fig. 2.

Representative chart records of two Hflour-loaded Xenopus single fibers during the two 2-min contractile fatigue periods in a blocked order. A: typical contraction curve under Po₂ of 3–5 Torr (first run) and Po₂ of 30 Torr (second run). B: typical contraction curve under Po₂ of 30 Torr (first run) and Po₂ of 3–5 Torr (second run).

Representative fluorescence of fibers in low Po₂ (3–5 Torr), as presented in Fig. 3A, resulted in an ablated signal due to ebselen treatment. However, during the second contractile period after ebselen washout, the signal was restored. Likewise, in high Po₂ (30 Torr), ebselen treatment was effective in eliminating the signal during the first contractile period, while fluorescence was restored after ebselen washout (see Fig. 3B).

Fig. 3.

Representative chart records of two Hflour-loaded Xenopus single fibers during the two 2-min contractile fatigue periods in a blocked order of ebselen treatment. A: typical contraction curve under Po₂ of 3–5 Torr with ebselen (first run) and Po₂ of 3–5 Torr (second run). B: typical contraction curve under Po₂ of 30 Torr with ebselen (first run) and Po₂ of 30 Torr (second run).

As illustrated in Fig. 4A, intracellular ROS formation during contractions was measured with confocal microscopy. In the low Po₂ condition, ROS levels were significantly elevated compared with high Po₂ (n = 10; P < 0.05). Ebselen scavenged ROS in both Po₂ conditions (n = 3; P < 0.05). Mean ROS levels over the first 120 s of intracellular ROS formation are shown during the four different treatments (Fig. 4B). ROS formation was significantly elevated 45 s after the initiation of contractions in low Po₂. However, in high Po₂, this increase was delayed for ∼30 s compared with low Po₂. In addition, ROS signals were ∼50% higher in low Po₂ compared with high Po₂ at the end of the contractile period (n = 10; P < 0.05). Note that the large dips in fluorescence curves (Figs. 2 and 3) do not represent actual ROS formation, but a motion artifact. To compensate for this, only the mean fluorescence values were used during the contractile period, so the random motion artifacts can be minimized (Fig. 4).

Fig. 4.

A: mean data for intracellular reactive oxygen species (ROS) generation during contractions with four different treatments in the single myofibers [Po₂ 30 Torr (n = 10), Po₂ 3–5 Torr (n = 10), Po₂ 30 Torr with ebselen (n = 3), and Po₂ 3–5 Torr with ebselen (n = 3)]. Fluorescence was recorded every 5 s during each contractile work period. *P < 0.05, Po₂ 3–5 Torr vs. Po₂ 3–5 Torr with ebselen. +P < 0.05, Po₂ 30 Torr vs. Po₂ 30 Torr with ebselen. #P < 0.05, Po₂ 3–5 Torr vs. Po₂ 30 Torr. B: mean data for intracellular ROS generation at the onset (focusing on the first 90 s of the contractile periods in A) among four different treatments as shown by changes in fluorescence in isolated single myofibers [Po₂ 30 Torr (n = 10), Po₂ 3–5 Torr (n = 10), Po₂ 30 Torr with ebselen (n = 3), and Po₂ 3–5 Torr with ebselen (n = 3)]. Fluorescence was recorded every 5 s during each contractile work period. *P < 0.05, Po₂ 3–5 Torr vs. Po₂ 3–5 Torr with ebselen. +P < 0.05, Po₂ 30 Torr vs. Po₂ 30 Torr with ebselen.

The grouped data show the effect of ebselen on muscle fatigue in Fig. 5. In high Po₂, the fall in force (fatigue) with ebselen treatment was not significantly different than without ebselen. However, the addition of ebselen in low Po₂ displayed significant protection against fatigue compared with high Po₂ with ebselen (Po₂ 3–5 Torr with ebselen, n = 3; Po₂ 30 Torr with ebselen, n = 3; P < 0.05; two panels on right). Furthermore, there was no significant difference in the initial maximal force and subsequent fatigue development among the following treatments: Po₂ 30 Torr (n = 10), Po₂ 3–5 Torr (n = 10), and Po₂ 30 Torr with ebselen (n = 3). Muscle viability was determined by calculating the mean specific initial force (mN/mm²) for each treatment group. These values are listed as follows: Po₂ 30 Torr, 1,993 ± 476 (n = 10); Po₂ 3–5 Torr, 2,396 ± 438 (n = 10); Po₂ 30 Torr with ebselen, 1,700 ± 388 (n = 3); and Po₂ 3–5 Torr with ebselen, 2,139 ± 116 (n = 3). There was no significant difference among these groups.

Fig. 5.

Data showing the decline in force (fatigue) during the four treatments in Xenopus single fibers [Po₂ 30 Torr (n = 10), Po₂ 3–5 Torr (n = 10), Po₂ 30 Torr with ebselen (n = 3), and Po₂ 3–5 Torr with ebselen (n = 3)]. *Significantly different from the initial force in each contractile period (P < 0.05). #Significantly different from Po₂ 30 Torr with ebselen (P < 0.05).

The maximal rate of increase of ROS fluorescence during contraction was calculated. During low Po₂, the rate was significantly higher than high Po₂ (1.529 ± 0.25 vs. 0.8540 ± 0.086 RU/min; n = 10, P < 0.05).

DISCUSSION

Major findings.

The present study demonstrates intracellular ROS formation in single frog myofibers during low Po₂ conditions. However, this oxygen level is normal for exercising human muscles (40). Former research has not been able to measure intracellular ROS formation during hypoxia in single contracting skeletal myofibers. This is because there is an O₂ diffusion gradient across the various layers of a nonperfused tissue, such as an entire rodent diaphragm. Therefore, it is difficult to precisely control hypoxic/normoxic conditions within the individual myofibers in a whole muscle preparation. It is likely that O₂ cannot completely penetrate the numerous layers of muscle tissue in whole muscle models, but instead hypoxic conditions predominate in the core of the muscle (1).

Previously, we have successfully measured ROS formation using hypoxic diaphragm muscle strips (46). However, a few issues exist regarding the whole muscle model: 1) A large population of vasculatures is present in skeletal muscle tissues, so it is difficult to determine whether ROS are generated from endothelium or myocytes. The advantage of using single myofiber systems is that we can locate the exact site where ROS are formed, as addressed in our previous study (36). 2) It is difficult to monitor the contraction of the whole muscle system under confocal microscopy because hundreds of moving cells in the system create a greater level motion artifact. However, it is easier to focus on one contracting single fiber with minimal motion effect using a confocal set-up. 3) To detect ROS formation or study the muscle physiology under an exact oxygen tension, such as Po₂ 3–5 Torr or Po₂ 30 Torr, a single myofiber preparation is an excellent model because there is no significant O₂ diffusion gradient across the various layers of the muscle tissue (36). Therefore, the current study overcomes these limitations by utilizing a single skeletal muscle fiber for reducing motion artifacts and minimizing oxygen gradients from extracellular to intracellular sources.

Although there is a lack of myoglobin in the Xenopus myofiber (24), myoglobin normally plays a role in maintaining oxygen homeostasis in mammalian muscle systems. Myoglobin may reduce any cellular damage from hypoxic stress. However, in our single myofiber model, we are generally more focused on the pure hypoxic effect on a single skeletal myofiber, since any myoglobin in our model could potentially cause a buffering effect, which may delay or interfere with the required low oxygen condition. In addition, although the frog fiber used in the current study lacks myoglobin, there is a great level of correlation between the frog and mammalian myocytes (9), and these two share a substantial similarity in typical skeletal muscle physiology, especially concerning mitochondrial and contractile function. Xenopus fibers are frequently used as a representative model for analyzing skeletal muscle function and metabolism (34, 47). Amphibian muscles demonstrate a close relationship between oxidative capacity and subsequent fatigue resistance (41). This effect is highly consistent with the corresponding skeletal muscle activity observed in mammals (41). In addition, mitochondrial function influences ROS activity in both amphibian and mammalian muscle fibers. Thus, although small differences between human (mammalian) and Xenopus myofibers exist, Xenopus single fibers provide an excellent model for studying an intracellular ROS response at high and low Po₂ conditions.

Importantly, our data showed that muscle stimulation caused a marked increase of the fluorescent signal during the low Po₂ condition. The change in the fluorescent signal was completely abolished by the addition of the antioxidant, confirming that our fluorescent signal was indicative of ROS generation. The data are consistent with previous evidence that suggested low levels of ROS formation occur in resting skeletal muscle and that signal strength is amplified during muscle contractions (7, 18, 29, 30). The present study, using real-time confocal imaging techniques, extends these findings from whole muscle to single myofibers, thereby supporting the observation that intracellular ROS formed within the myofiber is a generalized response to low Po₂ conditions (46).

ROS detection method.

Fluorescent detection from an intracellular probe sensitive to ROS is a common method used in studying the role of ROS formation in skeletal muscle. This is attributed to its high sensitivity for ROS detection at relatively low loading conditions (45). Among these probes, the most popular is dichlorodihydrofluorescein (DCFH), which is highly sensitive to H₂O₂ and is the most utilized probe for ROS detection in skeletal muscle systems (20, 29, 46). Dihydrofluorescein (Hfluor), a fluorescein analog of DCFH, has been found to have various superior characteristics to DCFH, including less photobleaching, higher molar fluorescence, lack of sensitivity to nitric oxide, and improved intracellular retention (10, 45, 46). Therefore, we used Hfluor in the current study to monitor ROS (H₂O₂) formation in real-time in intact single skeletal myofibers. Because Hfluor reacts with ROS, resulting in the formation of fluorescein (Fluor), we recorded the Fluor emission to monitor ROS. Confocal microscopy has some unique advantages compared with conventional fluorescence microscopy because it is more focused on the selected optical layer of interest. As a result, it significantly improves the imaging resolution in our system (44, 45).

Molecular sources of ROS generation in skeletal myofibers.

It has been suggested that there are a number of potential cellular sources of ROS in contracting skeletal muscle models. One possible source of elevated ROS formation in contracting fibers is from the mitochondria. ROS generation in mitochondria is believed to form by single-electron leakage to oxygen in the respiratory chain (4, 48). During respiration, most of the oxygen consumed during oxidative phosphorylation is reduced to water, while a minor portion (∼0.15%) of oxygen can be reduced to generate superoxide and hydrogen peroxide in pathological conditions (3, 33). Previous research (47) has shown that ROS are generated ∼15 s after the beginning of repetitive tetanic contractions when perfusate Po₂ is very high (∼160 Torr). However, in the present study, ROS fluorescence was significantly increased ∼75 s after the start of contractions at a Po₂ of 30 Torr and ∼45 s after the start of contractions under low Po₂ (3–5 Torr, Fig. 4). This suggests that the very high Po₂ (160 Torr) that we used previously stimulated a faster response for ROS formation compared with the Po₂ of 30 Torr and the low Po₂ of 3–5 Torr used in the present study (5, 14, 19). At 30 Torr, the activation of ROS takes the longest, likely because the intracellular antioxidant system is well preserved. During the low Po₂ condition, it is possible that ROS generation is slower than the high Po₂ condition (47) due to the low levels of oxygen (46). However, at the end of the 2-min contraction period in the very high Po₂ condition used in our previous study, ROS were elevated only ∼40% above baseline (47), while the low Po₂ condition of the present study resulted in ∼100% increase above baseline. Interestingly, at 30 Torr Po₂ in the present study, contraction-induced ROS fluorescence is similar to levels detected at Po₂ = 160 Torr in the previous study (47). Therefore, single myofibers initiate a quicker response during high Po₂ (∼15 s vs. 45 s) but do not accumulate as much ROS as is seen in low Po₂ conditions (∼40% vs. 100%). This may be due to a lower amount of overall oxidative stress associated with 30 Torr Po₂ conditions, but the more rapid response may be due to the very high levels of O₂ in the myofiber at the initiation of contractions.

Myoglobin has been associated with free radical generation under oxidative stress and can thereby cause skeletal muscle dysfunction (24). Since Xenopus skeletal muscle fibers do not contain myoglobin (11, 17), myoglobin is not a possible ROS generator in our model. There are additional sources of enzyme-based ROS formation that are likely stimulated during muscular activity. Particularly, NADPH oxidase (NOX) and xanthine oxidase (XO) have been shown to play a role in ROS formation during repetitive contractions and exhaustive exercise in skeletal muscle (13, 39). Since research has shown that skeletal muscle expresses NOX (35, 42), it could be one of the potential ROS generators, and we cannot rule that possibility out. However, some studies have suggested that ROS formation from NOX typically occurs in endothelial cells (23, 44). XO has typically been associated with more severe conditions of stress, such as exhaustive exercise (8, 39). It was unlikely in the current investigation that XO was the central source responsible for the detected ROS formation, since an intact single contracting myofiber retains no significant amount of vascular tissue on a dissected muscle fiber. Therefore, the most likely candidate responsible for ROS formation is the intracellular mitochondrion, although a detailed molecular mechanism of ROS development is still poorly understood in functional intact myofibers.

Kinetics of ROS generation.

ROS fluorescence was significantly elevated above baseline in both the low and high Po₂ conditions during tetanic contractions (Fig. 2). Fluorescent signals were greater during the low Po₂ condition because of additional ROS production (46). As illustrated in Fig. 2, the low and high Po₂ contractile treatments were independent of one another, demonstrating that there was no “preconditioning” effect observed in the current study. In addition, Fig. 4 shows that ROS generation became significantly elevated above baseline in both the low and high Po₂ contractile periods at 45 s and 75 s (P < 0.05), respectively, after the start of contractions. This 30-s additional time delay in the high-Po₂ condition was likely due to the endogenous antioxidant system being less overwhelmed by a lower oxidative stress within the myofiber, thus resulting in an extended period of time for significant ROS buffering.

It should be pointed out that the trend of ROS development varies at different O₂ levels. O₂^·− is the product of O₂ and free electrons donated by reducing molecules such as intra-mitochondrial NADH, which is a natural response to hypoxia in skeletal muscle (46) and single intact myofibers (12). During very high Po₂ (160 Torr), the excessive amount of the reactant O₂ could potentially trigger the early formation mechanism of O₂^·− compared with high Po₂ or low Po₂. However, because of the limited supply of free electrons under a Po₂ of 160 Torr (36), the accumulated ROS level was less than during the low Po₂ condition of the present study. At the Po₂ 30 Torr condition, which is in the regular physiological range for myofibers, both O₂ and free electrons remain relatively low, leading to slower (vs. high Po₂) and reduced (vs. low Po₂) ROS generation. In the low Po₂ condition of 3–5 Torr, skeletal myofibers contain abundant NADH and FADH₂ because of limited O₂ availability (12, 46), which ensures sufficient free electrons for potentially larger amounts of ROS formation. However, the intracellular O₂ level is extremely low, moderately delaying the response of O₂^·− formation (vs. higher Po₂). Interestingly, prolonged low Po₂ (>1 min) generates more free electrons in the mitochondria and thus creates more oxidative stress compared with other Po₂ conditions (47), as shown in Fig. 4. Therefore, the timing and production of ROS may be dependent on various levels of reducing agents and intramuscular Po₂.

In addition, the maximal rate of increase in fluorescence was significantly greater in the low vs. high Po₂ conditions in the present study, suggesting that the myofibers at low Po₂ underwent more oxidative stress, which induced a more rapid ROS burst than high Po₂. The biological significance of this is that the maximal rate of ROS formation in low Po₂ is likely related to overwhelming the endogenous antioxidant system.

Antioxidant protection during low Po₂ conditions.

It has been suggested that excessive skeletal muscle ROS formation diminishes muscle function capacity, whereas a low to moderate level of ROS is vital for redox signaling and normal muscle metabolism (22, 25, 29, 36). Oxidative stress disrupts the equilibrium between ROS and antioxidant (AOX) levels, specifically when ROS formation outperforms the buffering effect created by AOX therapy. As seen in Fig. 5, administration of ebselen in high Po₂ conditions did not significantly reduce muscle fatigue. However, in the low Po₂ condition, ebselen treatment was able to protect the muscle fiber from the deleterious effects of ROS. It is likely that AOX treatment was able to regulate excessive quantities of ROS within the more hypoxic muscle fiber, thereby allowing levels that did not further severely reduce muscle function. In the high Po₂ conditions, ebselen was not able to recover function and, as Fig. 3 demonstrates, abolished all traces of ROS generation. These interesting results suggest that AOX treatment may protect muscle function during low but not high Po₂ conditions. This indicates that muscle fatigue can be induced by multiple factors, with ROS generation playing a more prominent role in this process when the Po₂ is particularly low.

Ebselen is a cell-permeable GSH peroxidase mimic and utilizes GSH to remove intracellular ROS (31). Ebselen was shown to be more effective than N-acetylcysteine (NAC), particularly for scavenging ROS in the isolated skeletal muscle (46). To be specific, the major effect of this compound is to decrease H₂O₂ formation. It can also scavenge peroxynitrite under certain conditions (32). Similar to most drugs on the market, it does have certain side effects when used in overdosage. However, this was not the case in our experiment, since the proper dosage was carefully applied on the basis of previous literature (46, 47). Our preliminary data also showed that ebselen has no quenching effect on fluorescence signals, which is consistent with former research (46).

In an intramuscular environment, antioxidants protect against potential oxidative stress induced by normal muscle contractions. For instance, during exercise training, Powers et al. (28) observed increased activity of the antioxidant superoxide dismutase (SOD) in rat soleus muscle. Since skeletal muscle is composed of different fiber types, the antioxidant levels vary with each muscle on the basis of its function and oxidative capacity (15, 41). Fast glycolytic fibers, for example, display low levels of mitochondria and consequently, are not highly oxygen-dependent (15). As a result, antioxidant activity is relatively lower in these fibers compared with slow but more highly oxidative fibers, which exhibit enhanced antioxidant activity (16). Cell signaling pathways that respond to certain stresses, such as exercise, infection, or inflammation, regulate oxidant and antioxidant levels via ROS intermediates (15). Furthermore, minor levels of ROS have been shown to precondition the muscle for increased level of antioxidant expression, which alleviates severe oxidative stress that occurs later (27, 49). However, the exact physiological role ROS play during muscle homeostasis has still not been fully elucidated (27). Understanding the homeostasis of ROS and antioxidant interaction in skeletal muscle during various physiological conditions may shed light on new potential therapeutics to mitigate severe muscular oxidative stress.

Interestingly, we observed that hypoxia (3–5 Torr) per se had no effect on myofiber fatigue during the 2 min of contractions. In previous research, longer hypoxia periods (30 min) were used in the whole skeletal muscle system to achieve substantial hypoxic effect (21, 46). However, in the current study, we employed a brief hypoxic treatment (3–5 min) and only a 2-min contractile period to maintain myofiber integrity. During a short hypoxic period, although ROS could be formed from fatigue stress, it is likely that cellular antioxidants, such as SOD, are able to alleviate relatively mild oxidative stress. Thus, the key point here is that only 2-min period of contractions did not allow enough time for a difference in fatigue to occur between high and low Po₂ (21).

Perspectives and Significance

The results from the present study demonstrate that ROS are generated during tetanic contractions in Xenopus single intact myofibers at both low and high Po₂ conditions, but with higher levels of ROS production at low Po₂. The time it takes for intracellular ROS formation is not immediate with the initiation of contractions, and differs between high and low Po₂, likely resulting from the antioxidant buffering system being more rapidly overwhelmed when the Po₂ is reduced. Both high and low Po₂-induced ROS formation can be abolished by antioxidant ebselen treatment. Ebselen administration was shown to have a significant effect in protecting myofibers contracting in low Po₂ conditions against fatigue development. Conversely, under high Po₂ conditions, ebselen was unable to rescue endogenous muscle function. This suggests that the mechanisms inducing fatigue development in low vs. high Po₂ conditions are likely different, with ROS generation playing a more prominent role when intracellular Po₂ is reduced. Thus, we speculate that intramuscular antioxidant defense is more active to alleviate muscle dysfunction under a more hypoxic condition.

GRANTS

This study was supported by National Heart, Lung, and Blood Institute Grant PPG-HL-091830, National Institute of Arthritis and Musculoskeletal and Skin Diseases Grant AR040155, Ohio State University College of Medicine Startup Funding 013000, OU General Fund G110, and Research Excellence Fund of Biomedical Research.

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the authors.

AUTHOR CONTRIBUTIONS

Author contributions: L.Z., P.D.W., and M.C.H. conception and design of research; L.Z. and A.S. performed experiments; L.Z. and W.J.R. analyzed data; L.Z., W.J.R., M.T.C., P.D.W., and M.C.H. interpreted results of experiments; L.Z. and W.J.R. prepared figures; L.Z., W.J.R., and M.T.C. drafted manuscript; L.Z., W.J.R., M.T.C., P.D.W., and M.C.H. edited and revised manuscript; L.Z., A.S., W.J.R., M.T.C., P.D.W., and M.C.H. approved final version of manuscript.